1Lung Biology Laboratory, Division of Pulmonary, Allergy, and Critical Care Medicine, Department of Medicine, Columbia University College of Physicians and Surgeons, New York, New York, USA.
2Department of Orthopedic Surgery, School of Medicine, Keio University, Tokyo, Japan.
3Department of Pediatrics, Columbia University College of Physicians and Surgeons, New York, New York, USA.

First published April 25, 2011Submitted: May 26,
2010; Accepted: February 9,
2011.

Shedding of the extracellular domain of cytokine receptors allows the diffusion of soluble receptors into the extracellular space; these then bind and neutralize their cytokine ligands, thus dampening inflammatory responses. The molecular mechanisms that control this process, and the extent to which shedding regulates cytokine-induced microvascular inflammation, are not well defined. Here, we used real-time confocal microscopy of mouse lung microvascular endothelium to demonstrate that mitochondria are key regulators of this process. The proinflammatory cytokine soluble TNF-α (sTNF-α) increased mitochondrial Ca2+, and the purinergic receptor P2Y2 prolonged the response. Concomitantly, the proinflammatory receptor TNF-α receptor–1 (TNFR1) was shed from the endothelial surface. Inhibiting the mitochondrial Ca2+ increase blocked the shedding and augmented inflammation, as denoted by increases in endothelial expression of the leukocyte adhesion receptor E-selectin and in microvascular leukocyte recruitment. The shedding was also blocked in microvessels after knockdown of a complex III component and after mitochondria-targeted catalase overexpression. Endothelial deletion of the TNF-α converting enzyme (TACE) prevented the TNF-α receptor shedding response, which suggests that exposure of microvascular endothelium to sTNF-α induced a Ca2+-dependent increase of mitochondrial H2O2 that caused TNFR1 shedding through TACE activation. These findings provide what we believe to be the first evidence that endothelial mitochondria regulate TNFR1 shedding and thereby determine the severity of sTNF-α–induced microvascular inflammation.

Introduction

Although inflammation initiates as a defensive response against pathogenic stimuli, progression to severe inflammation leads to tissue injury. In the short term, severe inflammation causes life-threatening edema in critical organs such as the lung and brain. In the longer term, inflammation associates with multiple diseases including vascular plaque formation in atherosclerosis, malignant transformation, chronic obstructive pulmonary disease, and fibrotic diseases of the lung, liver, and kidney (1–5). There is therefore considerable interest in understanding mechanisms that regulate progression of the inflammatory process from defense to injury.

Ectodomain shedding of cytokines and cytokine receptors plays a major role in establishing this balance, as exemplified by the cytokine receptor TNF-α receptor–1 (TNFR1), which is critical to inflammatory progression (6, 7). The process initiates with TNFR1 ligation by soluble TNF-α (sTNF-α), plasma levels of which rise during inflammation (8, 9). A signaling cascade results, leading to NF-κB activation and proinflammatory gene transcription (10) as well as activation of the cell surface metalloprotease TNF-α converting enzyme (TACE; ref. 11). Also referred to as adisintegrin and metallopeptidase domain 17 (ADAM17; ref. 12), TACE cleaves TNFR1. The TNFR1 ectodomains released in the extracellular space chelate sTNF-α, providing negative feedback to the TNF-α–induced inflammatory loop (13). Pathogenic bacteria, including Chlamydia trachomatis, Neisseria meningitides, and Staphylococcus aureus, have evolved TNFR1 shedding mechanisms as a strategy for circumventing the host immune response (14–16).

The role of mitochondria in these inflammatory events requires consideration. sTNF-α increases production of mitochondrial H2O2 (17) that could activate TACE by oxidizing cysteine thiols in the TACE prodomain (18). Mitochondrial superoxide dismutase catalyzes the dismutation of superoxide produced in the electron transport chain (ETC) to H2O2. The site of superoxide production in the ETC depends on the cell type, being predominantly at complex I in skeletal muscle and neural cells (19), but at complex III for vascular smooth muscle cells (20), ECs (21), and alveolar epithelial cells (22). Mitochondrial H2O2 diffuses across the mitochondrial outer membrane to access cytosolic targets, causing multiple functional outcomes, including mechano-oxidative coupling (23), insulin resistance (24), stabilization of hypoxia-inducible factor–1α (20), and tissue factor production (25). However, the role of mitochondrial H2O2 in endothelial TNFR1 shedding remains unclear.

The vascular endothelium is a major regulator of inflammation. Many acute inflammatory diseases result from inflammation in microvascular beds, in which endothelial activation causes transcriptional expression of leukocyte adhesion receptors such as E-selectin (26). Endothelia of both large and small blood vessels express TNFR1 (27), which is critical to this process, as indicated by the fact that inflammation is abrogated in mice lacking TNFR1 (6). Endothelial TNFR1 shedding has been demonstrated in cultured human umbilical vein ECs (28) and in ECs incubated with microparticles from atherosclerotic plaques (29). However, it is not known whether microvascular endothelium in situ sheds TNFR1 and whether the shedding regulates the inflammatory process.

In myeloid cells, TNFR1 shedding commences in minutes (30), which suggests the involvement of rapidly activated signaling mechanisms attributable to second messengers such as Ca2+. The optically imaged lung provides an excellent platform for in situ studies of endothelial second messengers (17, 23). We took advantage of this approach to determine the role of endothelial Ca2+ as a determinant of endothelial TNFR1 shedding in microvessels. We tested this hypothesis by exposing lung microvascular endothelium to blood levels of sTNF-α occurring in sepsis (8, 9). Our findings indicate that sTNF-α causes endothelial TNFR1 ectodomain shedding through release of mitochondrial H2O2, implicating mitochondria as regulators of the inflammatory response.

Results

Receptor shedding. To detect surface expression of TNFR1 on ECs in situ, we determined microvascular immunofluorescence using mAb MCA2350, which binds extracellular TNFR1. TNFR1 fluorescence colocalized with an endothelial cytosolic marker (Figure 1A) and was stable for at least 1 hour (data not shown). The membrane-impermeable fluorescence quencher trypan blue (TB) abolished the fluorescence (Figure 1A), which affirmed that the immunofluorescence was on the endothelial surface. A 10-minute infusion of human sTNF-α initiated fluorescence loss that continued well after the end of sTNF-α infusion (Figure 1C) and was concentration dependent (Supplemental Figure 1A; supplemental material available online with this article; doi:
10.1172/JCI43839DS1). Buffer infusion was without effect (Figure 1B). Coinfusion of mAb E20, which blocks TNF-α binding to TNFR1 (31), inhibited the fluorescence loss (Figure 1D), ruling out nonspecific effects. Responses caused by TNF-α receptor 2 (TNFR2) can be ruled out, since mAb E20 is specific for TNFR1, and human sTNF-α does not ligate mouse TNFR2 (32).

We considered the role of TACE, the proteolytic enzyme that cleaves surface TNFR1 on leukocytes (11). Since it is not known whether TACE cleaves TNFR1 in lung microvessels, we used the EC-specific TACE-deleted (EC-Tace–/–) mouse (33). sTNF-α–induced TNFR1 shedding was completely blocked in EC-Tace–/– mice, but not in Tacefl/fl floxed mice (Figure 1E). TNF-α–processing inhibitor–1 (TAPI-1), which inhibits TACE, also inhibited sTNF-α–induced shedding (Figure 1E). These findings affirmed that sTNF-α–induced fluorescence loss resulted from TNFR1 shedding through activation of endothelial TACE.

We determined microvascular immunofluorescence of TNFR1 in a mouse model of pneumonitis established by instilling intratracheal LPS. In WT mice, but not in Tnfa–/– mice, LPS markedly decreased microvascular TNFR1 expression compared with intratracheal instillation of control buffer (Figure 1F). These findings affirm that LPS caused TNF-α–dependent, microvascular TNFR1 shedding.

To determine global lung responses, we gave 10-minute intravascular infusions of sTNF-α through the main pulmonary artery. After 30 minutes, we homogenized the lungs and performed IB on lysates of lung tissue using mAb E20 that recognizes cytosolic TNFR1. Shedding of the TNFR1 ectodomain leaves a 27-kDa cytosolic fragment (34). The appearance of this fragment in WT and Tacefl/fl mice, but not in Tace–/– mice, confirmed that the receptor’s ectodomain was shed. The continued presence of the 55-kDa band in sTNF-α–treated lungs (Figure 1G) indicated that nonendothelial TNFR1 — as, for example, that expressed in alveoli (31) — was not shed, probably because the infused sTNF-α did not access TNFR1 at all lung sites. Flow cytometry of ECs, identified as the vWF-positive fraction in primary cell isolates from lung vessels, revealed loss of TNFR1 (Figure 1, H and I), affirming that the shedding response occurred in the lung as a whole.

Internalization versus shedding. To distinguish between receptor internalization and receptor shedding, each of which could underlie the loss of TNFR1 fluorescence, we determined endothelial uptake of the fluorescent TNFR1 ligand Alexa Fluor 488–conjugated sTNF-α (sTNF-α–488). We normalized fluorescence of sTNF-α–488 against the cytosolic fluorescence of calcein red. As a comparison, we determined uptake of FITC-conjugated albumin (FITC-albumin), which is internalized by endothelium (35). As expected, infusion of FITC-albumin increased green fluorescence of the endothelial lining (Figure 2, A and B). Failure of TB to diminish the fluorescence (Figure 2, B, C, and G) confirmed that the infused FITC-albumin was taken up by the endothelium. In contrast, the immediate postinfusion fluorescence of sTNF-α–488 on the endothelial lining was quenched by TB (Figure 2, D and E), confirming binding of the ligand to extracellular TNFR1. However, sTNF-α–488 did not colocalize with calcein (Figure 2F), which indicates that the ligand was not internalized. sTNF-α–488 fluorescence decreased with a time course similar to that of TNFR1 fluorescence (Supplemental Figure 1, B and C). Together, these findings indicate that ligated TNFR1 was shed and not internalized.

A consistent feature was the prolonged duration of the Ca2+ responses that persisted for up to 1 hour, lasting well beyond the 10-minute sTNF-α infusion. Since increase of mitochondrial Ca2+ increases ATP production by the ETC (39), and since ATP is secreted (40), we considered the role of the purinergic receptor P2Y2 (encoded by P2Y2). In P2Y2–/– mice, both cytosolic and mitochondrial Ca2+ oscillations were augmented during sTNF-α infusion, as in WT mice. However, the prolonged oscillations were completely inhibited (Figure 4). In contrast, no inhibition occurred in P2Y1 receptor–deficient P2Y1–/– mice (Figure 4), which indicates that the P2Y2 receptor was responsible for the prolonged Ca2+ responses.

To determine the mitochondrial role in ROS production, we infused microvessels with the mitochondria-specific antioxidant MitoQ (41), which consists of the lipophilic triphenylphosphonium (TPP) cation bound to the antioxidant ubiquinol. The negative mitochondrial potential localizes the cationic compound to mitochondria, where the MitoQ complex undergoes oxidation reduction reactions. In MitoQ-treated microvessels, the sTNF-α–induced decrease of roGFP fluorescence was completely blocked (Figure 5, B and C), which indicates that MitoQ blocked ROS production. As a positive control, we confirmed that exposure to H2O2 decreased the fluorescence (Figure 5, B and C). The inhibition by MitoQ indicated that the ROS were of mitochondrial origin.

In microvessels in which we detected ROS by the dichlorofluorescein (DCF) method, the mitochondrial role was further affirmed: the sTNF-α–induced DCF response was completely blocked by pretreating vessels with the mitochondrial complex I inhibitor rotenone (Supplemental Figure 4, A and B). The ROS response was not inhibited in microvessels treated with the flavoprotein inhibitor diphenylene iodonium or the eNOS antagonist rotenone (G)-nitro-l-arginine methyl ester (l-NAME), which indicated that the DCF responses were specific to ROS. Further, the ROS response was not inhibited in microvessels of Nox2–/– mice (Supplemental Figure 4C), ruling out a role for this NOX isoform.

To identify the ROS, we overexpressed the H2O2 hydrolyzing agent catalase targeted to mitochondria and the cytosol (mCAT and cCAT, respectively; Supplemental Figure 5, A–D). We used DCF fluorescence as the ROS reporter. In microvessels expressing these catalases, the sTNF-α–induced ROS increases were completely inhibited (Figure 5D), whereas no inhibition occurred with the empty vector (data not shown). The inhibition by mCAT indicated that ROS fluorescence was caused by mitochondrial H2O2. The inhibition by cCAT indicated that H2O2 diffused from mitochondria to the cytosol. The similar inhibitions of ROS by mCAT and cCAT indicate that overexpression of catalase at either site was sufficient to reduce all of the H2O2 in the cell. In microvessels of WT mice treated with MitoQ or expressing mCAT or cCAT, TNFR1 shedding was abrogated, whereas shedding was not blocked in microvessels of Nox2–/– mice (Figure 5E). sTNF-α–induced mitochondrial Ca2+ increases were not affected by MitoQ treatment or mCAT overexpression (Supplemental Figure 3), ruling out the possibility that these treatments blocked the sTNF-α–induced mitochondrial Ca2+ response.

The Rieske iron sulphur protein (RISP) of mitochondrial complex III is a major source of mitochondrial ROS in lung cells (20). To determine the role of RISP in the present responses, we intravenously injected mice with fluorophore-labeled siRNA against RISP (siRISP) complexed with cationic liposomes. These procedures did not increase endogenous sTNF-α in blood or in the BAL, and leukocyte BAL counts were unchanged (Supplemental Figure 6, B and C), affirming that the liposome-siRNA treatment did not cause detectable innate immune responses (42). At 2 days after injection, in mitochondrial fractions of lung ECs or lung tissue lysates, RISP was markedly lower in siRISP-treated lungs than in those treated with scrambled siRNA (scRNA; Figure 6, A and B, and Supplemental Figure 6A). Although a faint RISP band was present in the nonmitochondrial membrane fraction (Figure 6B), there was no knockdown of this band, which suggests it was caused by nonspecific binding by the IB antibody. No RISP band was present in the cytosolic fraction (Figure 6B). In isolated lung mitochondria, RISP knockdown decreased rates of both ATP production and O2 consumption (Figure 6, C and D, and Supplemental Figure 6D). The mitochondrial uptake of a potential-sensitive dye was similar in siRISP- and scRNA-treated lungs (Supplemental Figure 6E), consistent with reports that the present ATP decrease does not change the mitochondrial membrane potential (43).

siRISP fluorescence in microvessels was patchy: fluorescence was bright in some regions, but relatively dim in others (Figure 6E). After sTNF-α infusion, siRISP-expressing microvessels showed sustained TNFR1 fluorescence (Figure 6, F and H), which indicates that RISP knockdown abrogated the shedding response. In contrast, the infusion induced loss of TNFR1 fluorescence in scRNA-expressing microvessels (Figure 6, G and H), which indicates that shedding was present in these vessels. We interpret that knockdown of mitochondrial RISP protected against TNFR1 shedding; this is the first evidence to our knowledge that mitochondrial RISP determines ectodomain shedding of a proinflammatory receptor.

In one experiment, we took advantage of the uneven siRISP expression in different microvessels in order to determine whether blockade of TNFR1 shedding was also uneven. At baseline, there was no correlation between siRISP and TNFR1 fluorescence (Supplemental Figure 7A), which indicates that RISP knockdown did not affect the expression of unligated TNFR1. In contrast, sTNF-α infusion induced a strongly positive correlation between the expression levels of RISP and TNFR1 (Supplemental Figure 7B), which indicates that sites of high siRISP expression were also sites at which TNFR1 shedding was blocked. These findings affirm that the extent of mitochondrial RISP expression determined the extent of TNFR1 shedding.

To assay TACE expression in the alveolo-capillary region, we determined immunofluorescence of the TACE extracellular domain by injecting fluorescent antibodies by alveolar or microvascular micropuncture. At baseline, TACE immunofluorescence was well developed on both alveolar epithelium and microvascular endothelium in Tacefl/fl mice (Figure 7A). However, in EC-Tace–/– mice, TACE fluorescence was absent on the endothelium, although VE-cadherin fluorescence (positive control) was present (Figure 7A). In EC-Tace–/– mice or in microvessels given TAPI-1 infusion, the sTNF-α–induced cytosolic and mitochondrial Ca2+ and ROS responses were enhanced (Figure 7, B–D). These findings indicate that sustained expression of TNFR1, resulting from inhibition of TNFR1 shedding, caused enhanced sTNF-α–induced Ca2+ and ROS responses. We interpret these findings to suggest that ligation of TNFR1 by sTNF-α led to TACE activation by mitochondria-derived H2O2.

Leukocyte recruitment. To assess inflammatory responses, we determined microvascular expression of the leukocyte adhesion receptor E-selectin. Although not present at baseline, E-selectin fluorescence was clearly evident 2 hours after the 10-minute sTNF-α infusion (Figure 8, A and B). This response was diminished in P2Y2–/– mice (Figure 8D), which indicates that the P2Y2 receptor played a role in inducing endothelial E-selectin. The sTNF-α infusion also increased microvascular leukocyte adhesion in 2 hours (Figure 9, A–C). Both E-selectin expression and leukocyte adhesion responses were markedly enhanced in microvessels treated with siRISP or with inhibitors of mitochondrial electron transport (Figure 8, A–D, and Figure 9C). These findings indicate that inhibitors of mitochondrial function blocked the inflammatory responses.

Discussion

sTNF-α caused rapid TNFR1 shedding in lung microvessels, providing the first evidence to our knowledge that ECs in situ shed the receptor. Injection of sTNF-α also increased endothelial cytosolic Ca2+, mitochondrial Ca2+, and mitochondrial ROS. These responses confirm our previous findings in which IP3-induced increase of cytosolic Ca2+ sequentially caused the mitochondrial responses, independently of external Ca2+ (17, 23). We now report that TNFR1 shedding was blocked after inhibition of mitochondrial Ca2+ by RR or inhibition of ROS by mitochondrial RISP knockdown. Overexpression of mCAT, which hydrolyzes H2O2, also blocked the shedding. These findings indicate that H2O2 is the ROS responsible for the shedding and that mitochondrial Ca2+ increase is upstream of H2O2 production. Together, our previous (17, 23) and present findings suggest that mitochondrial Ca2+ increase caused RISP-induced ROS production in microvessels, leading to endothelial TNFR1 shedding.

Our results indicate that an inflammatory response occurred despite TNFR1 shedding. Thus, the sTNF-α–treated microvessels expressed endothelial E-selectin. Induction of E-selectin, which is not normally expressed by microvessels, indicates that sTNF-α caused proinflammatory transcription (44, 45) that might have been induced by the increase in cytosolic Ca2+ (46, 47). Microvascular leukocyte recruitment was also evident, occurring possibly as a consequence of increased E-selectin expression. Notably, when we blocked mitochondrial Ca2+ and ROS production, TNFR1 shedding was blocked. Consequently, endothelial TNFR1 expression was sustained, resulting in a stronger inflammatory response. This was indicated by enhanced E-selectin expression and leukocyte recruitment. To our knowledge, these findings provide the first evidence for a signaling sequence in which mitochondrial inhibition enhances the inflammatory response to sTNF-α (Figure 10). We propose that TNFR1 shedding by mitochondrial H2O2 protects against progression of inflammation.

The Ca2+ increases occurred in 2 phases: first as an initial phase during the 10-minute sTNF-α infusion, and then as a prolonged phase that persisted for up to 1 hour, even after the infusion had stopped. Increase of mitochondrial Ca2+ activates mitochondrial dehydrogenases, increasing ATP production by the ETC (37). ECs secrete ATP, which ligates the P2Y2 receptor to induce IP3-dependent cytosolic Ca2+ increases (40, 48). Here, lack of the P2Y2 receptor, namely in P2Y2–/– mice, blocked the prolonged phase of Ca2+ oscillations, but not the initial phase. Concomitantly, the induced E-selectin expression was also markedly suppressed. Although the P2Y2 receptor has been implicated in leukocyte recruitment (49), to our knowledge, underlying signaling mechanisms have not been defined. Our findings indicate that by prolonging microvascular Ca2+ oscillations, the P2Y2 receptor critically regulates the inflammatory response.

Endothelial TACE (12) has previously been implicated in pathological neovascularization (33). Our findings are suggestive of a new role for endothelial TACE, namely as a determinant of the microvascular inflammatory response. Under inflammatory conditions in which leukocytes adhere to ECs, endothelial ecto­domain shedding could potentially be activated by proteases on the leukocyte surface (50). Definitive exclusion of this possibility was achieved by the availability of mice carrying floxed alleles of Tace and a Cre recombinase expressed in ECs (Tie2-Cre). Although mice lacking TACE die in the perinatal period (51), EC-Tace–/– mice survive to adulthood, expressing no obvious developmental abnormalities (33). In these adult EC-Tace–/– mice, there was no sTNF-α–induced TNFR1 shedding, which indicates that the shedding was entirely regulated by endothelial TACE.

TACE activity is enhanced by oxidation of cysteine thiol groups in its disintegrin/cysteine-rich region (18). A proposed mechanism of TACE activation is that proinflammatory second messengers such as ROS (52, 53) and NO (54) disrupt the thiol-zinc linkage, resulting in shedding of TACE substrates such as TNFR1 (55). Consistent with this proposal, we found that after infusion of MitoQ (41) or overexpression of cCAT or mCAT, TNFR1 shedding was blocked. These findings, taken together with the considerations discussed above, indicate that sTNF-α–induced mitochondrial H2O2 activated TACE to establish the shedding responses.

Loss of TNFR1 from the cell surface could be due to shedding or cellular internalization. Cultured ECs grown in static media internalize shed TNFR1 ectodomains (56). However, in an in situ assay that we believe to be novel, we failed to detect any evidence for endothelial internalization of the TNFR1 ectodomains, although our assay successfully detected albumin internalization, thereby ruling out nonspecific errors. Extracellular release of TNFR1 ecto­domains was also indicated in flow cytometry and IB analyses of primary vascular cell isolates recovered from sTNF-α–treated lungs. We conclude from these findings that in the presence of blood flow, which is likely to wash out cleaved proteins from the endothelial surface, shed ectodomains are not internalized by microvascular endothelium.

Redox studies in vivo have been hampered by nonspecificities attributable to pharmacological inhibitors and fluorescein-based dyes. In cultured cells, the ROS-sensitive protein roGFP provides specific quantification of cytosolic H2O2 (57). In its first in vivo application to our knowledge, we expressed the roGFP-encoding construct in microvessels. In roGFP, engineered cysteines form a disulfide crosslink between β strands adjacent to the chromophore in GFP (58). Specificity of the probe as an oxidant reporter is attributable to the fact that cysteine oxidation causes structural, hence spectral, changes. Our validation studies confirmed that roGFP was a reliable indicator of H2O2 for in situ determinations. In addition, we selectively overexpressed catalase in the mitochondria or the cytosol. These approaches, including the RISP knockdown with siRNA, provided specific strategies for addressing mitochondrial redox responses in vivo.

The clinical significance of our findings bears on the emerging importance of mitochondria in inflammatory disease. Here, we show that TNFR1 shedding, an antiinflammatory effect, occurred in a TNF-α–dependent manner in lung microvessels as well as in LPS-treated lungs. To the extent that endothelial mitochondria are responsible for the shedding response, as we show here, loss of mitochondrial function might be proinflammatory. Inflammatory conditions such as sepsis cause loss of lung mitochondrial function, as demonstrated by decreased lung ATP (59, 60). Our findings indicate that in such conditions, therapy aimed at restoring endothelial mitochondrial function might protect against the deleterious effects of increasing inflammation.

Animal procedures were approved by the Institutional Animal Care and Use Committee of Columbia University Medical Center. Tacefl/fl mice and EC-Tace–/– mice (Tacefl/flTie2-Cre; ref. 33) were provided by C.P. Blobel (Hospital for Special Surgery, Medical College of Cornell University, New York, New York, USA). P2Y1–/– and P2Y2–/– mice (both C57BL/6 background) were provided by B.H. Koller (University of North Carolina, Chapel Hill, North Carolina, USA). Nox2–/– and Tnfa–/– mice (both C57BL/6 background) and age-, gender- and strain-matched WT mice were purchased from Jackson Laboratory.

Lung preparation

Lungs were prepared according to our previously reported methods (61). Briefly, mice were anesthetized (ketamine-xylazine), and lungs were excised en bloc. Lungs were continuously pump-perfused with autologous blood (final hematocrit, 10%) diluted in HEPES (150 mmol/l Na+, 5 mmol/l K+, 1.0 mmol/l Ca2+, 1 mmol/l Mg2+, and 20 mmol/l HEPES at pH 7.4) containing 4% dextran (70 kDa) and 1% fetal bovine serum at pH 7.4 and osmolarity of 295 mosM at a rate of 0.5 ml/min at 37°C. Lungs were constantly inflated through an airway cannula at airway pressure of 5 cmH2O. Pulmonary artery and left atrial pressures were held at 10 and 3 cmH2O, respectively. The lung surface was moistened with saline and covered with Saran Wrap during imaging to prevent drying. For assay of leukocyte adhesion, we enriched the lung perfusate with 2.5 × 105 cells/ml of fluorescently labeled (calcein red) leukocytes (62) for 30 minutes followed buffer wash for 20 minutes to remove nonadherent leukocytes. Leukocytes were counted in each lobe of the lung from images taken with a ×10 objective.

Imaging

Live fluorescence imaging was carried out using our previously described methods (23). Briefly, microvessels were fluorescence loaded by 20-minute infusions of fluorophores given in the lung perfusion, followed by a 10-minute buffer wash. Microvessels were viewed by confocal microscopy (LSM 5; Zeiss). Ca2+ responses were recorded at 1 image/10 seconds and DCF at 1 image/30 seconds. The longer interval prevented dye photoactivation. To rule out potential errors attributable to local changes in cell volume, fluorescence of Ca2+ and ROS-sensitive dyes was normalized against fluorescence of a soluble cytosolic dye (calcein or calcein red). In all experiments in which more than 1 dye was loaded, we confirmed absence of bleed-through between fluorescence emission channels. Spatial selectivity of cytosolic (fluo-4 AM) and mitochondrial (rhod-2 AM) Ca2+ dyes was determined by infusion of saponin at the end of an experiment, which selectively abolishes cytosolic Ca2+ fluorescence (63). To assay fluorophore internalization, infusion of fluorophore-conjugated agents was followed by vascular infusion of the fluorescence-quenching agent TB. To label endothelial surface proteins (TNFR1 and E-selectin), fluorophore-conjugated antibodies were infused by micropuncture (64) of lung microvessels for 5 minutes followed by buffer wash to remove unbound antibody. Endothelial fluorescence was quantified using MetaMorph software through analyses based on region-of-interest (Ca2+ and ROS), line scan (antibody fluorescence), and segmentation (leukocyte adhesion) tools.

Results are expressed as mean ± SEM. 1-way ANOVA followed by Newman-Keuls post-hoc analyses (MiniTab) was used for comparisons between groups. A P value less than 0.05 was accepted as significant. All group data were obtained from experiments repeated in separate lungs unless otherwise stated.

Supplemental data

Acknowledgments

This work was supported by NIH grants HL57556, HL36024, and HL69514 to J. Bhattacharya. The TNF-α ELISAs were supervised by C.W. Schindler (Columbia University Medical Center, New York, New York, USA). Tara Guclu assisted with manuscript preparation.

Footnotes

Conflict of interest: The authors have declared that no conflict of interest exists.